Regulation of DNA Replication within the
Immunoglobulin Heavy-Chain Locus During B Cell
Commitment
Agnieszka Demczuk1, Michel G. Gauthier2, Ingrid Veras1, Settapong Kosiyatrakul3, Carl L. Schildkraut3,
Meinrad Busslinger4, John Bechhoefer2, Paolo Norio1,5*
1 Department of Oncology, Montefiore Medical Center, Moses Division, Bronx, New York, United States of America, 2 Department of Physics, Simon Fraser University,
Burnaby, Canada, 3 Department of Cell Biology, Albert Einstein College of Medicine, Bronx, New York, United States of America, 4 Research Institute of Molecular
Pathology, Vienna Biocenter, Vienna, Austria, 5 Department of Medicine, Albert Einstein College of Medicine, Bronx, New York, United States of America
Abstract
The temporal order of replication of mammalian chromosomes appears to be linked to their functional organization, but the
process that establishes and modifies this order during cell differentiation remains largely unknown. Here, we studied how
the replication of the Igh locus initiates, progresses, and terminates in bone marrow pro-B cells undergoing B cell
commitment. We show that many aspects of DNA replication can be quantitatively explained by a mechanism involving the
stochastic firing of origins (across the S phase and the Igh locus) and extensive variations in their firing rate (along the locus).
The firing rate of origins shows a high degree of coordination across Igh domains that span tens to hundreds of kilobases, a
phenomenon not observed in simple eukaryotes. Differences in domain sizes and firing rates determine the temporal order
of replication. During B cell commitment, the expression of the B-cell-specific factor Pax5 sharply alters the temporal order
of replication by modifying the rate of origin firing within various Igh domains (particularly those containing Pax5 binding
sites). We propose that, within the Igh CH-39RR domain, Pax5 is responsible for both establishing and maintaining high rates
of origin firing, mostly by controlling events downstream of the assembly of pre-replication complexes.
Citation: Demczuk A, Gauthier MG, Veras I, Kosiyatrakul S, Schildkraut CL, et al. (2012) Regulation of DNA Replication within the Immunoglobulin Heavy-Chain
Locus During B Cell Commitment. PLoS Biol 10(7): e1001360. doi:10.1371/journal.pbio.1001360
Academic Editor: Tom Misteli, National Cancer Institute, United States of America
Received January 25, 2012; Accepted May 30, 2012; Published July 10, 2012
Copyright: ! 2012 Demczuk et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits
unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was funded by NIH-NIGMS grant R01GM080606 to PN. MGG and JB are funded by HSFP and NSERC-Canada. CLS and SK are funded by
R01GM045751 and ESSCF–NYS–C024348. MB is funded by Boehringer Ingelheim. The funders had no role in study design, data collection and analysis, decision to
publish, or preparation of the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
Abbreviations: 4-OHT, 4-hydroxy-tamoxifen; CldU, 59-chloro-29-deoxyuridine; IAS, number of initiation events occurring within a specific section of the locus,
per allele, per S phase; IdU, 59-iodo-29-deoxyuridine; pre-RC, pre-replication complex; SMARD, single molecule analysis of replicated DNA.
* E-mail: paolo.norio@einstein.yu.edu
Answering these questions requires a quantitative understanding
of the dynamics of origin firing. Based on measurements of
average origin activity across entire genomes, various stochastic
models of origin firing have been recently used to explain specific
aspects of eukaryotic DNA replication, such as the duration of S
phase [6–13]. If origin firing can occur stochastically anywhere
along the genome and at any time during S phase, origin
distribution and the timing of origin firing cannot be responsible
for establishing the temporal order of replication [14]. Recent
observations indicate that the profile of replication timing of the
budding yeast genome can be explained by differences in the firing
rate of individual origins and stochastic origin firing [15].
However, yeast differs from metazoans in many aspect of DNA
replication (e.g., S. cerevisiae has well-defined origins of replication,
lacks the developmental control of the temporal order of
replication, shows no correlation between gene expression and
the temporal order of replication, has a short S phase, etc.). In
addition, previous studies have mostly relied on the measurement
of individual parameters of DNA replication which can be
modeled with limited detail to determine the dynamics of origin
firing (e.g., the timing of replication) and can produce misleading
results if applied to complex genomes [16]. Hence, testing this
Introduction
During the S phase, mammalian chromosomes replicate in a
precise temporal order, with the timing of replication typically
changing gradually across hundreds of kilobases. Cell differentiation induces regional changes in the order of replication which
can affect 45%, or more, of the mouse genome [1]. Various studies
have examined how the temporal order of replication is established and modified at specific gene loci, but provided
discordant explanations about the role played by DNA origins of
replication. For example, within a 340 kb portion of the Igh locus,
changes in replication timing have been linked to modifications in
the distribution of active origins and in their firing efficiency (see
definitions in Table 1) [2]. In contrast, within the beta-globin locus,
changes in replication timing can occur without significant
changes in origin distribution, or firing efficiency, and have been
ascribed to modifications in the timing of origin firing [3–5]. Does
this mean that the temporal order of replication is determined by
multiple mechanisms? Are origin distribution, firing efficiency, and
the timing of origin firing regulated independently? Which aspect
of origin activation is controlled by cell differentiation? These are
some of the questions addressed in this study.
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Developmental Control of Origin Activity
and firing rates that determines the temporal order of replication.
This organization remains valid for cells at different stages of B cell
development (e.g., bone marrow pro-B cells blocked at the
uncommitted and committed stages of differentiation by homozygous mutations of the Pax5 and Rag2 genes; Figure 1B). We also
show that the changes in DNA replication that occur during B cell
commitment can be quantitatively explained by substantial
changes in the firing rate of origins within specific domains of
the Igh locus. Therefore, the rate of origin firing is the parameter
that is being regulated across large sections of the locus during cell
differentiation. The role of the developmental regulator Pax5 in
this process and its mechanism of action are also examined.
Author Summary
Each time a mammalian cell duplicates its genome in
preparation for cell division it activates thousands of so
called ‘‘DNA origins of replication.’’ The timely and
complete duplication of the genome depends on careful
orchestration of origin activation, which is modified when
cells differentiate to perform a specific function. We
currently lack a universally accepted model of origin
regulation that can explain the replication dynamics in
complex eukaryotes. Here, we studied the mouse immunoglobulin heavy-chain locus, one of the antibodyencoding portions of the genome, where origins change
activity when antibody-producing B cells differentiate in
the bone marrow. We show that multiple aspects of DNA
replication initiation, progression, and termination can be
explained mathematically by the interplay between randomly firing origins and two independent variables: the
speed of progression of replication forks and the firing rate
of origins along the locus. The rate of origin firing varies
extensively along the locus during B cell differentiation
and, thus, is a dominant factor in establishing the temporal
order of replication. A differentiation factor called Pax5 can
alter the temporal order of replication by modifying the
rate of origin firing across various parts of the locus.
Results
In Uncommitted Pro-B Cells the Replication of the Igh
Locus Follows a Precise Temporal Order of Replication
Bone marrow pro-B cells isolated from Pax52/2Rag22/2 mice
retain the ability to proliferate but are blocked at the uncommitted
stage of differentiation and maintain the Igh locus in germline
configuration [20]. In order to perform SMARD, we sequentially
labeled a population of exponentially growing cells with 59-iodo29-deoxyuridine (IdU) and 59-chloro-29-deoxyuridine (CldU), for
3–4 h (see Table S1, column b). Under these conditions, each
labeling period is long enough to allow the complete replication of
large sections of the Igh locus, resulting in DNA molecules variably
substituted with the halogenated nucleotide (see example in Figure
S1) [2,17,21,22]. We then digested the genomic DNA with
restriction enzymes that cut infrequently within the locus (PmeI, or
PacI, or SwaI) and isolated four of the resulting restriction
fragments by pulsed-field gel electrophoresis (gray bars in
Figure 1A). These fragments were stretched on microscope slides,
hybridized with specific DNA probes, and analyzed by fluorescence microscopy to detect the incorporation of the nucleotide
analogs along individual DNA molecules.
The molecules fully substituted with halogenated nucleotides
provide a vast amount of information about the process of DNA
replication [2,17,21,22]. In Pax52/2Rag22/2 cells, the analysis of
,4,000 hybridization signals yielded 1,304 fully substituted DNA
molecules, 764 of which met the standards required to perform
precise measurements (Table S1, columns c–f). This population
includes molecules that incorporated only one type of halogenated
nucleotide (single-labeled molecules), as well as molecules that
incorporated both of them (double-labeled DNA molecules). As
explained in previous publications [17,22], the ratio between singleand double-labeled molecules is proportional to the time required to
replicate each restriction fragment (Table S1), which is linked to the
average number of replication forks participating in the replication
of each restriction fragment and to their average speed (Table S1).
Thus, these values can be directly determined from the experimental data as described in the legend of Table S1. This allowed us to
determine that, in uncommitted pro-B cells, the average speed of
replication forks is similar within the four restriction fragments
(between 2.25 and 3 kb/min). A similar value was also measured at
the genomic level using a different assay (,2.5 kb/min; IV and PN,
unpublished observation). This suggests that replication forks move
at comparable speeds throughout the Igh locus.
In the double-labeled DNA molecules, IdU-CldU transitions
mark the positions of replication forks at the time of the label
switch. Initiation events appear as IdU-labeled regions surrounded
by CldU, while fork collisions display the complementary pattern
(e.g., Figure 1C–D). Figure 2A summarizes the location of each
initiation event (red bars) and fork collision (green bars) detected in
158 individual restriction fragments. The normalized frequency of
hypothesis rigorously in mammalian cells requires the measurement of multiple parameters of DNA replication.
Any large portion of a mammalian genome can be used to test
the stochastic firing of origins provided enough information is
available about DNA replication. For this reason, we used the
assay called single molecule analysis of replicated DNA (SMARD)
[17] to collect unbiased information about all aspects of DNA
replication initiation, progression, and termination across a 1.4
megabase region encompassing the mouse immunoglobulin
heavy-chain (Igh) locus (Figure 1A). The experimental data sets
collected by SMARD included the temporal order of replication,
the steady-state distribution of replication forks, the time required
to replicate the region, the average speed of replication forks, the
distribution of initiation and termination events, the percentage of
replicating molecules containing initiation and termination events,
and the average number of events per molecule.
Using a novel mathematical procedure [18], we established that
the experimental data sets collected by SMARD are fully
consistent with the stochastic firing of origins as defined in
Table 1. We also show that many aspects of DNA replication
(including the temporal order of replication) can be explained by
variations in the rate of origin firing I(x,t) along the Igh locus (see
definition in Table 1). According to the nomenclature proposed by
others [19], this rate indicates the number of initiation events
occurring per length of unreplicated DNA, over a given period of
time, as mathematically defined in Materials and Methods.
Our results point to significant differences in the regulation of
DNA replication between the mouse Igh locus and yeast
chromosomes. In S. cerevisiae, each origin of replication appears
to be characterized by a specific firing rate, which differs for
different origins [15]. Within the Igh locus, initiation events lack
focal points corresponding to individual origins. Instead, the locus
comprises large domains (spanning tens or hundreds of kilobases)
where the firing rate of multiple origins is virtually uniform and
similarly regulated. Along the locus, changes in the firing rate
occur abruptly at the border between different Igh domains, and
the firing rate of each domain is not affected by deletions that span
multiple origins. It is the combined effect of different domain sizes
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Table 1. Definitions for various terms used in the text.
Origin of Replication
Any genomic site at which DNA replication can begin. In some organisms, these sites can be identified from their DNA sequence; in others
they cannot.
Origin Density
The average number of distinct potential origin of replication present within a genomic regions of specified size in a defined population of
cells. This definition implies that replication complexes must form at each of these sites in at least a fraction of the cell population (but not
necessarily at every potential origin in each individual cell of the population).
Origin Firing
Activation of bidirectional replication forks at a potential origin.
Passive Replication
Replication of a potential origin by a fork generated elsewhere.
Origin Efficiency
The parameter most frequently used to indicate the level of origin activity (e.g., the relative intensity of bubble arcs in neutral-neutral 2D-gel
experiments). For a genomic region of defined size and location, it represents the ratio between the number of firing events and the total
number of replication events (firing events + passive replication). In other words, it expresses the frequency at which origins fire within the
region during each S phase. Since this value is affected by external events (causing the passive replication of the region), it is not an intrinsic
property of origins.
When a genomic region contains a single origin, efficiency is always a value between 0 and 1. For example, a value of 0.5 indicates that
during each S phase a specific origin fires in 50% of the cells.
In contrast, if the region contains multiple origins the value is always $0. In the article, we refer to this latter value with the acronym IAS
(initiations per allele per S phase).
Timing of Origin Firing A common interpretation of the replication dynamics in eukaryotic cells assumes that specific origins fire at specific times during the S phase.
Stochastic Origin
Firing
The model used for the analysis of our experimental data is stochastic in that it assumes that potentially active origins are present at high
density throughout the Igh locus and that firing can take place in any unreplicated portion of the locus at any time during the S phase. The
rate at which origins fire along the locus can vary and is determined by the cellular context.
Origin Firing Rate
For an unreplicated portion of the genome of defined size and location, the firing rate indicates the number of origin firings per unit of time
that occur as cells transit through the S phase. This value measures an intrinsic property of origins within specific portions of the genome,
which persists from the time of the G1/S transition, through the S phase, until the region is fully replicated.
For example, a value of 50061026 initiations/kb/min indicates that, within a specific 1 kb section of the genome, 500 origins will fire every
minute in a population of 106 unreplicated DNA molecules (alleles).
doi:10.1371/journal.pbio.1001360.t001
the dynamics present at the steady-state of growth (right panel in
Figure S1) [2]. This means that the population of double-labeled
DNA molecules can be used to determine the steady-state
distribution of replication forks across the Igh locus (arrowheads
in Figure 2B–C), and the average number of forks participating in
the replication of each restriction fragment (Table S1, column l).
Similarly, the temporal order of replication can be obtained from
the level of IdU substitution of these molecules, with peaks
marking the regions that replicate first and valleys marking the
regions that replicate last (Figure 2D). Overall, these results
indicate that, in uncommitted pro-B cells, DNA replication tends
to begin near the DH-JH and middle-VH gene families, as well as at
origins located 39 and 59 of the Igh locus. From there, replication
forks proceed to replicate the locus until they collide with an
oppositely moving fork (predominantly within the termination
regions).
the events across the four fragments is also indicated (expressed as
number of events scored per 100 double-labeled DNA molecules,
per 100 kb). The values before normalization are presented in
Table S1. Note that initiation events and fork collisions spanning
adjacent restriction fragments are not shown in the figure since
these events are scored as simple IdU-CldU transitions (a fact that
is taken into consideration by our mathematical model and in
Figure S2C). Overall, these results indicate that initiation events
are more frequent near the DH-JH and middle-VH gene families
than in other parts of the locus. In these regions, the presence of
non-overlapping events implies the existence of clusters of active
origins that are reminiscent of initiation zone [23,24]. In contrast,
fork collisions are more frequent between the origin clusters
(termination regions in Figure 2A).
It is important to point out that SMARD experiments are
designed to detect only a fraction of the active origins (the number of
double-labeled DNA molecules required to reach mapping
saturation increases with the number of potential origins).
Moreover, reaching mapping saturation is increasingly unlikely
where origin efficiency is low, and near the end of the restriction
fragments. Thus, Figures 2A and S2 are expected to largely
underestimate the actual density of origins along the locus (see
definition in Table 1). Even so, the distance between the midpoints
of initiation event within the DH-JH region suggests an origin density
higher than one potential origin per 10 kb (Figure S2E, PacI#3). In
Figure 2A, origin density appears lower near the middle-VH region
genes. However, within this section of the locus, the midpoints of
initiation events overlap only rarely, indicating that that origin
mapping is far from saturation (Figure S2E, PmeI#5). Lack of
saturation is even more pronounced across the proximal-VH region
(none of the detected initiation events overlap; Figure 2A, PmeI#4).
We conclude that potential origins of replication are likely to be
present at a relatively high density across most of the Igh locus.
Since cells grow asynchronously during DNA labeling, the IdUCldU transitions depict all stages of DNA replication, reflecting
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The Replication Dynamics of the Igh Locus Are Consistent
with the Stochastic Firing of Origins and with Variations
of Their Firing Rate along the Genome
The data presented above show that the replication of the Igh
locus follows a precise temporal program. However, this program
represents a population average. At the level of individual DNA
molecules, initiation events do not seem to occur in any particular
order, with different origins firing in different molecules. Initiation
events were also detected within the portions of the Igh locus that
replicate last (e.g., the termination region within PmeI#4 and the
region between Cd and Cc3; Figure 2A), taking place as the
corresponding molecules were at various stages of their replication.
Such events are not easily explained by deterministic models of
origin activation. A domino activation of origin firing [13] also
seems unlikely. The initiation events detected in the population of
double-labeled DNA molecules are rarely associated with externally generated replication forks, and when such forks are present
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Developmental Control of Origin Activity
Figure 1. Experimental system and approach used in this study. (A) Map of the Igh locus (129/Sv) with approximate positions of the various
gene families—constant (CH), joining (JH), diversity (DH), proximal-variable (proximal-VH; VH7183, VHQ52), middle-variable (middle-VH; VHS107), distalvariable (distal-VH; VHJ606, VHJ558, VH3609P)—and regulatory elements—intronic enhancer (Em), and 39 regulatory region (39RR). Gray bars indicate
the positions of the four restriction fragments analyzed in this study. Note that while PmeI #5 and PmeI #4 span a continuous portion of the locus, a
gap of ,20 kb is present between the right end of PmeI #4 and the left end of PacI #3. A short overlap is also present between PacI #3 and SwaI
#2. (B) Schematic representation of B cell development. Rag2 mediates the step of DNA cleavage during DH-JH and VH-DJH recombination. When this
gene is mutated, cells cannot assemble the pre-B cell receptor and are unable to develop beyond the committed pro-B stage of differentiation. In
contrast, Pax5 regulates the expression of hundreds of genes involved in B cell commitment, and in its absence, cell differentiation stops at the
uncommitted pro-B cell stage. (C) SMARD. a–f represent examples of staining patterns for six hypothetical double-labeled DNA molecules. Diverging
and converging forks indicate the occurrence of initiation and termination events. (D) Images of 12 actual molecules representative of the categories
depicted in panel C. Each image is aligned to the map of the corresponding genomic fragment using the position of ‘‘landmark’’ hybridization probes
as a reference. Arrowheads mark the position of the IdU-CldU transitions.
doi:10.1371/journal.pbio.1001360.g001
throughout the Igh locus and the S phase (see Table1 for a precise
definition), generating forks that move at a constant speed. In these
calculations, the rate of origin firing was the only parameter
allowed to vary freely across the Igh locus, while remaining
constant in time (meaning that, within a genomic region, initiation
events continue to occur at the same rate from the beginning of S
phase until the region is replicated in the entire population of
cells). It is important to point out that the rigid constrains imposed
by this scenario do not accurately reflect physiological conditions.
For example, it is known that modest changes in the speed of
replication forks can indeed occur along the genome and during
the S phase [25]. However, by limiting the number of free
variables, this scenario allows us test the stochastic firing
hypothesis more stringently.
Strikingly, we found that this simple scenario is sufficient to
reproduce all experimental data sets collected by SMARD
their location is tens to hundreds of kilobases from the firing
origins (e.g., 40 and 100 kb in the example shown in Figure S2A).
Hence, it is possible that Igh origins fire according to stochastic
dynamics.
To determine whether the results obtained by SMARD can be
quantitatively explained by stochastic origin firing (see definition in
Table 1), we used a mathematical formalism and a simulation
procedure that we recently developed for this purpose [18]. As
briefly described in Materials and Methods, this procedure allows
us to fit many of the data collected by SMARD (namely, the
distribution of the replication forks, Figure 2B–C, the temporal
order of replication, Figure 2D, and the replication time of the
restriction fragments, Tr, Figure S2B) to computer-generated
curves calculated from a series of rate equations.
In the simplest possible scenario, the curves were calculated
assuming that bidirectional origins of replication fire stochastically
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Figure 2. Replication of the Igh locus in Pax52/2Rag22/2 uncommitted pro-B cells (129/Sv). The panels containing experimental data (A–D)
appear discontinuous because independent experiments were performed for each restriction fragment. (A) The top portion of this panel shows the
schematics of the four restriction fragments depicted in Figure 1A (shown to scale). Red and green bars indicate the positions of the initiation and
termination events detected by SMARD. Numerals indicate the normalized frequency of the events scored in each population of restriction fragments
(this value represents an underestimate of the actual number of events taking place during every replication cycle, a fact that is taken in
consideration by our mathematical model; see Materials and Methods). These events are organized from top to bottom by increasing size of the
labeled region. Some bars are shown adjacent to each other to emphasize that they occupy non-overlapping locations; this does not mean that the
events occurred on the same DNA molecule. The midpoint on each bar is marked by black symbols to indicate the most likely position of an active
origin or fork collision. Gray arrows show the predominant directions of replication fork movement across the region. (B, C) Distribution of the
replication forks moving leftward and rightward, from SMARD (empty symbols), and from the fit (line). Each value indicates the number of replication
forks per kb (fork density) detected in the population of double-labeled DNA molecules after averaging the result over intervals of defined size (10 kb
for the PmeI fragments, 5 kb for PacI#3 and SwaI#2). Trends in fork direction can be detected by comparing, for individual fragments, the densities
shown in (B) and (C). The densities of different fragments cannot be compared directly because this value is a function of the replication time, which
varies for each fragment (Table S1, Column i). Error bars were calculated as described in Materials and Methods. (D) IdU content of the population of
double-labeled DNA molecules averaged over intervals of defined size (see panel B), obtained experimentally by SMARD (empty symbols), and from
the fit (line). For each of the four restriction fragments, individual data points indicate the fraction of double-labeled molecules substituted with IdU
within that interval. Since each fragment was studied individually, discontinuities are visible. However, adjacent fragments show similar trends in the
shape of their graphs (increasing or decreasing), allowing us to deduce the position of the regions replicating first (peaks) and last (valleys). Error bars
were calculated as described in Materials and Methods. (E) Distribution of the firing rate of origins across the Igh locus (blue line) and at adjacent
genomic locations (red and blue arrows). Changes in the firing rate are shown continuously along the 1.4 megabase region spanned by the four
restriction fragments. The firing rate is expressed in initiations/kb/minute to indicate that during each minute of the S phase, within a specific 1 kb
section of the locus, a certain number of origins will fire in a population of 1026 unreplicated DNA molecules (see also Table 1). This figure also shows
the efficiency of origin firing within specific sections of the Igh locus (gray dashed lines), with gray numerals indicating the number of initiation events
occurring in each region, per allele, per S phase (IAS). (F) Curves showing extent of DNA replication at different times in S phase for the various parts
of the Igh locus (10%, 50%, and 90% replicated).
doi:10.1371/journal.pbio.1001360.g002
Another striking feature of these results is that the firing rate
changes abruptly at a few specific locations (transitions), while
remaining virtually uniform across large sections of the locus
(plateaus). Attempts at fitting bell-shaped curves (which lack both
plateaus and sharp transitions) produced fits of lower quality
compared to box-shaped curves of the kind shown in Figure 2E
(M.G.G., J.B., and P.N., unpublished observation). Therefore, the
Igh locus appears to be organized in precisely defined domains
where origins have a similar rate of firing (suggesting a significant
degree of coordination among the origins of individual domains).
We conclude that the temporal order of replication is a
consequence of the combined effect of domain sizes and firing
rates.
Equation 3, in Materials and Methods, also allows us to
determine the efficiency of origin firing for specific sections of the
Igh locus, according to the definition provided in Table 1. The
values obtained for the DH-JH and middle-VH regions are only 1.1
and 1.5 initiation events, per allele, per S phase (IAS, gray dashed
lines in Figure 2E). These regions are known to contain multiple
origins of replication (Figure 2A). Hence, most of the origins
remain silent during each cycle of replication (firing is inefficient).
Notably, the portion of Igh locus spanning the proximal-VH genes
produces 0.4 IAS even if its firing rate is about 2 orders of
magnitude lower than the DH-JH and middle-VH regions. The
nonlinear relationship between firing rate and origin efficiency can
be explained by the fact that the proximal-VH genes occupy a very
large genomic region and tend to replicate after adjacent portions
of the locus (Figure 2D). This provides more time for the origins
therein located to fire. Hence, efficiency is not an intrinsic property
of individual origins, but rather the result of the distribution of the
firing rate throughout the Igh locus.
The computer-generated data set allows us to draw a few
additional conclusions. Although Igh origins fire inefficiently, they
are responsible for the replication of 87% of the locus, with only
minor contributions from external origins (Figure S2F). This
means that inefficient origin firing extends beyond the margins of
the locus into the surrounding regions. In addition, various
portions of the locus have a very high probability of being
replicated by forks moving in one particular direction (peaks and
valleys in Figure S2G). Since the molecules analyzed in this
(Figures 2B–D and S2B–C). This scenario can even reproduce
data sets that were not used during the fitting procedure (such as
the location of initiation events and fork collisions, the number of
molecules containing such events, the average number of events
per molecules, and the average speed of replication forks). Since
the fit was performed simultaneously for all restriction fragments,
the presence of local discrepancies is not particularly surprising
(e.g., PmeI #4 in Figures 2D and S2B). This is likely to reflect the
constrains imposed by this scenario (e.g., within some portions of
the locus fork speed may deviate from the average and origin
density may also vary). Nevertheless, the high quality of the fit
(reduced chi-square, 1.18) indicates a close match between
calculated and experimental data sets. Hence, the results collected
by SMARD are fully compatible with the stochastic firing of
origins throughout the Igh locus.
In addition to the changes in firing rate along the locus,
scenarios involving a larger number of variables were also considered (e.g., allowing for a variable speed of replication forks or
for changes in the rate of origin firing during the S phase). In
principle, changing these parameters could have a major effect on
the replication dynamics of the Igh locus. However, these scenarios
improved the quality of the fit only marginally (M.G.G., J.B., and
P.N., unpublished observation). This means that variations in
parameters other than the firing rate of origins along the genome,
while possible, have a limited impact on the replication dynamics
and the temporal order of replication of the Igh locus in uncommitted pro-B cells.
The Temporal Order of Replication Is Determined by
Wide Differences in the Firing Rate of Origins across
Large Sections of the Igh Locus
The high quality of the fit calculated above indicates that the
computer-generated data set provides a good approximation of the
firing rate of origins across the Igh locus. Figures 2E and S2D show
that the firing rate is very low throughout most of the locus
(4.561026 initiation events per kb per minute). Two exceptions
are the DH-JH region (55613.2 kb in size) and of the middle-VH
region (281643.1 kb in size), where the firing rate is up to 77-fold
higher. This variation implies that origins located in different parts
of the Igh locus have a very different tendency to fire.
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Figure 3. Replication of the Igh locus in a DH-JH rearranged clonal population of Pax52/2 uncommitted pro-B cells (129/Sv-C57BL/6).
The results for each allele are shown separately. Only two restriction fragments were analyzed (PacI #3 and SwaI #2). (A–F) Summary of the results
obtained by SMARD from the analysis of 1,443 hybridization signals, yielding 532 fully substituted DNA molecules, 313 meeting the standards of
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measurement, for a total of 67 double-labeled molecules (see Table S1 for details). The results of the fitting procedure are shown as described for
Figure 2.
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experiment originated from both Igh alleles, the strong bias in fork
direction indicates that both alleles follow a similar replication
program. Finally, we calculate that it takes approximately 4 h to
complete the replication of this portion of the genome in 100% of
the cell population (Figure 2F). Hence, these experiments provide
a complete description of DNA replication, within the Igh locus,
over a broad portion of the S phase.
(Figure 3E). This is reminiscent of results obtained by studying
deletions of the DHFR locus by 2D-gel electrophoresis (although
in that case conclusions were based on measurement of origin
efficiency, which is context dependent, and not of firing rate)
[26,27]. We conclude that firing rate of origins is regulated
independently in different sections of the locus and that origin
activity at one location is not affected by the presence, or absence,
of neighboring origins.
The Firing Rate Is Regulated Independently in Different
Domains of the Igh Locus
B Cell Commitment Alters the Firing Rate of Origins
throughout the Igh Locus with the Largest Changes
Occurring in Domains that Contain Pax5 Binding Sites
Bone marrow pro-B cells from Pax52/2 mice are blocked at the
uncommitted stage of differentiation but, unlike Pax52/2Rag22/2
cells, they can undergo DH-JH recombination. The resulting loss of
various sections of the DH-JH origin cluster (the ,55613.2 kb
region containing the origins with the highest firing rate; Figure 2E)
provides us with a tool to study how the firing rate is regulated.
Specifically, we can use ex vivo propagation to obtain clonal
populations of cells carrying particular DH-JH deletions. Here, we
studied a heterozygous pro-B cell clone that carries a 65 kb
deletion on its 129/Sv allele and a 25 kb deletion on the C57BL/6
allele. The different sizes of the resulting restriction fragments
allowed us to use SMARD to investigate the effect of each deletion
on DNA replication.
Consistent with the loss of the entire DH-JH origin cluster, the
65 kb deletion reduced the number of initiation events occurring
within fragment PacI#3 to an undetectable level (left portion of
Figure 3A and Figure S3B). The direction of replication fork
movement (Figure 3B–C) and the uniform slope of the temporal
order of replication (Figure 3D) indicate the passive replication of
the region by forks originating 39 of the Igh locus. This is similar to
results previously obtained in non-B cells, where the Igh locus is
part of a replication timing transition region [2]. Besides for the
loss of the DH-JH origins, the rate of origin firing in adjacent
portions of the locus remains unchanged (compare PacI#3 and
SwaI#2 in Figures 2E and 3A). This rate is very low compared to
the origins located at 39 of the Igh locus. Hence, even if the 59-end
of PacI#3 now replicates 3 h later compared to unrearranged proB cells (compare Figure 3F at 280 kb to the corresponding portion
of Figure 2F), this delay is not sufficient to produce large numbers
of initiation events within PacI#3 and SwaI#2 (infrequent
initiation events may still occur within this portion of the Igh
locus but their visualization would require the analysis of a much
larger sample of double-labeled DNA molecules).
In contrast, the 25 kb deletion removes less than half of the DHJH origin cluster. On this allele, initiation events and fork collisions
continue to occur at the same locations described for Igh locus in
germline configuration (compare the right portion of Figure 3A
with Figure 2A). However, there is a strong right-to-left bias in the
direction of fork movement (Figure 3B–C), and the IdU content
shows a steady decrease in the same direction with an inflection
point at the site of the remaining DH-JH origins (Figure 3D). These
factors indicate the passive replication of the region in a fraction of
the cell population. Accordingly, there is a 40% reduction in the
number of initiation events occurring within this section of the
locus during each S phase (0.7 IAS, Figure 3E), compared to the
level detected in unrearranged pro-B cells (1.1 IAS, Figure 2E).
These results indicate that the decrease in initiation events is
proportional to the reduction in size of the DH-JH origin cluster.
Hence, the firing rate of the remaining Igh origins and the location
of the firing rate transitions are unaffected by the 25 kb deletion
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In order to study how the replication of the Igh locus changes
during cell differentiation, we isolated B cell progenitors from the
bone marrow of a Rag22/2 mouse (129/Vs.). These cells efficiently undergo B cell commitment but maintain the Igh locus in
germline configuration, which prevents them from developing any
further. In these cells, initiation events and fork collisions are
distributed across most of the Igh locus (Figure 4A). Only a few of
these initiation events are centered at the same genomic location
(Figure S4), indicating that the experiment is far from mapping
saturation (leading to an underestimate of origin density). This
suggests that potentially active origins are present at high density
across most of the Igh locus, perhaps every 10–20 kb, although
larger gaps may exist at a few locations. Despite the widespread
activation of origins, it is still possible to distinguish portions of the
locus where replication forks have a preferred direction of
movement (e.g., PacI#3; Figure 4B–C) and regions that replicate
first and last (Figure 4D). Thus, our results indicate that the level of
origin activity is not uniform across the Igh locus and point to
major differences in DNA replication between committed and
uncommitted pro-B cells.
Once more, we found that the data obtained by SMARD are
fully compatible with the stochastic firing of origins (reduced chisquare, 1.03; Figure S4C). The nearly perfect fit confirms that the
temporal order of replication is mostly determined by the
regulation of a single variable along the locus (the firing rate of
origins). Computer calculations also show that the rate of origin
firing is not uniform (blue line in Figure 4E). A comparison with
the results obtained for uncommitted pro-B cells (red dotted line)
reveals that B cell commitment is accompanied by changes in the
rate of origin firing across most of the Igh locus. However, the
largest changes (whether positive or negative) occur in regions
where a previous genomic screening identified the presence of
Pax5 binding sites in committed pro-B cells [28]. For example, the
firing rate increases nearly 50-fold across the CH-39RR region
(,216 kb in size and containing two Pax5 binding sites) and
decreases 10-fold throughout the DH-JH origin cluster (,55 kb in
size and containing one Pax5 binding site). These results indicate
that the rate of origin firing is the parameter that is being regulated
during cell differentiation. In addition, they suggest that Pax5
participates in regulating the firing rate of origins during B cell
commitment (although the number of known binding sites for this
factor is far smaller than the number of potential origins affected
by its expression).
As a result of B cell commitment, changes in origin efficiency
were also observed throughout the Igh locus (compare gray dotted
lines in Figures 2E and 4E). However, the overall efficiency of
origins across the 1.4 Mb region increases only from 3.2 IAS in
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Figure 4. Replication of the Igh locus in Rag22/2 committed pro-B cells (129/Sv). (A–F) Summary of the results obtained by SMARD from the
analysis of 973 fully substituted DNA molecules, 683 meeting the standards of measurement, for a total of 213 double-labeled molecules (see Table
S1 for details). The results of the fitting procedure are shown as described for Figure 2. The locations of Pax5 binding sites in Rag22/2 committed proB cells, as well as some chromatin modifications, are also indicated in panel E (see Materials and Methods for additional details).
doi:10.1371/journal.pbio.1001360.g004
the firing rate of Igh origins that are similar to those observed in
pro-B cells isolated from mice.
Additional experiments confirmed that these effects are specific.
For example, the firing rate of CH-39RR origins does not increase
when 4-OHT is provided to uncommitted pro-B cells that do not
express Pax5ER (Table S1). In addition, the induction of Pax5ER
in KO-Pax5ER pro-B cells does not affect origin activity broadly
and non-specifically. For example, we did not observe significant
changes in cell proliferation and cell-cycle profile (Figure S5L) or
the appearance of markers of DNA damage and DNA damage
checkpoint activation (Figure S5M). We conclude that the change
in origin activity observed in KO-Pax5ER pro-B cells results from
the activation of Pax5ER and the induction of B cell commitment.
uncommitted pro-B cells (Figure S2D) to 7.4 IAS in uncommitted
pro-Bs (Figure S4C). Thus, the firing rate can increase by 1–2
orders of magnitude across most of the Igh locus but produce
variations of only 2.3-fold in the total number of initiation events
(and replication complexes) involved in the replication of the locus.
Given the high density of potential origins throughout the Igh
locus, these results also suggest that the firing efficiency of
individual origins is below 10%. Therefore, following B cell
commitment, origins continue to fire inefficiently even if the firing
rate increases across most of the Igh locus.
Inducing Pax5ER Increases the Firing Rate of Origins
within the CH-39RR Region and Changes the Temporal
Order of Replication
The Regulation of Firing Rate of Origins Does Not Require
Changes in Gene Expression and Takes Place in Late G1
Reconstituting Pax5 expression in Pax52/2 pro-B cells induces
B-cell commitment [20,29] and can be used to study the role of
this protein in origin regulation. For this purpose, we transduced
bone marrow pro-B cells from a Pax52/2 mouse (129/Sv-C57BL/
6) with a retroviral vector containing the expression cassette
Pax5ER-IRES-GFP [30,31]. We then sorted and expanded the
GFP+ cells to obtain a polyclonal population expressing Pax5ER
(KO-Pax5ER pro-B cells; Figure S5A–B). This protein is the
fusion product of Pax5 and the hormone-binding domain of the
estrogen receptor, which becomes biologically active in the
presence of 4-hydroxy-tamoxifen (4-OHT). In KO-Pax5ER proB cells, 4-OHT is able to induce commitment-specific changes in
the methylation of the 39RR DNA, indicating that Pax5ER is able
to interact with at least one of the Pax5 binding sites of the CH39RR region [32]. In KO-Pax5ER pro-B cells, the occurrence of
B-cell commitment was monitored by the Pax5-dependent
expression of the cell-surface-marker CD19. Before the addition
of 4-OHT, we consistently found that only 4% of KO-Pax5ER
pro-B cells are CD19+ (Figures 5A and S5C). However, 65%–90%
of cells become CD19+ after the addition of 4-OHT. Thus, in our
inducible system, the activity of Pax5ER is modest in the absence
of 4-OHT but increases dramatically after induction, leading to B
cell commitment.
Can the induction of Pax5ER increase the firing rate of origins
within the CH-39RR region? To answer this question, we studied
KO-Pax5ER pro-B cells before and after induction with 4-OHT
for 28 h (see labeling scheme in Figure 5B). This induction time
was chosen because it allows enough time for the synthesis and
turnover of the gene products regulated by Pax5 and for cells to
become fully committed to the B lineage [30]. In the absence of 4OHT, we detected a limited number of initiation events within the
CH-39RR region, which is consistent with the modest activation of
Pax5ER described above (first and third column in Figure 5C).
However, this portion of the locus continues to be passively
replicated in the majority of cells, as indicated by the replication
fork distribution and the temporal order of replication (Figures 5D–
F and S5H–I). In contrast, 4-OHT induction profoundly alters all
aspects of DNA replication within the CH-39RR region, suggesting
a strong increase in origin activity (second and fourth column in
Figures 5C–F and S5H–I). Numerical calculations confirm that
the firing rate of origins reaches the level detected in committed
pro-B cells (compare the 129/Sv results in Figures 5G and 4E).
Hence, inducing B cell commitment in vitro produces changes in
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B cell commitment involves changes in the expression of
hundreds of tissue-specific genes [33], raising the question of
whether Pax5 is directly responsible for regulating the firing rate of
origins. After the addition of 4-OHT, Pax5ER requires only a few
minutes to translocate from the cytoplasm to the nucleus, where it
changes the expression of its target genes over a period of several
hours [30]. We suspected that the firing rate would change very
slowly if its regulation depended on the activation or silencing of
these genes. Therefore, we decided to determine the speed at
which the firing rate increases within the CH-39RR region
following the addition of 4-OHT to KO-Pax5ER pro-B cells.
The labeling scheme for the time-course experiment is
presented in Figure 6A. Our experimental design takes advantage
of the fact that SMARD detects the occurrence of initiation events
in the population double-labeled DNA molecules. These molecules begin their replication during a precise interval of time Tr
that preceded the label switch, meaning that they originate from
cells that entered S phase sometimes before each molecule started
replicating [2]. As a result, we can study how DNA replication
initiates, progresses, and terminates across the genomic region of
interest, knowing that these events occurred in cells that crossed
the G1/S transition during a specific window of time from the
addition of 4-OHT (this procedure also avoids cell synchronization
and experimental manipulations that may generate experimental
artifacts).
Surprisingly, we found that the replication of the CH-39RR
region changes very rapidly after the addition of 4-OHT. For all
induction times between 1 and 24 h, the distributions of replication forks and the temporal orders of replication are similar to
the experiment performed at 28 h (compare Figure 6B–E with
Figure 5D–F). To accurately determine how the firing rate
changes over time, we simultaneously fit all experimental data sets
obtained for the 129/Sv allele (including those presented in
Figure 5). This resulted in a reduced chi-square of 1.02 (Figure
S6A–C). As summarized in Figure 6H, the firing rate of the CH39RR origins reaches a level comparable to committed pro-B cells
within the first hour of induction. Given that this induction time is
similar to the replication time of the restriction fragment
(approximately 1 h; Table S1), we conclude that the firing rate
of origins began to increase within minutes after the addition of 4OHT. This rapidity implies that the changes in firing rate do not
depend on modifications in gene expression and are likely to be
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Figure 5. Replication of the CH-39RR region in KO-Pax5ER pro-B cells (129/Sv-C57BL/6). (A) Kinetics of expression of CD19 after 4-OHT
induction. The number of CD19+ cells reaches a plateau at about 30 h. (B) Scheme of Pax5ER induction and DNA labeling used for the SMARD. (C–H)
Summary of the SMARD experiments performed to study the replication of CH-39RR region before and after induction with 4-OHT (from the analysis
of 1,832 fully substituted DNA molecules, 1,445 meeting the standards of measurement, for a total of 339 double-labeled molecules; Table S1). Each
experiment is presented in a different column. Results for the two Igh alleles are shown separately (129/Sv, C57BL/6). The results of the fitting
procedure are shown as described for Figure 2.
doi:10.1371/journal.pbio.1001360.g005
firing within the CH-39RR region. We then isolated the population
of committed pro-B cells (CD19+ fraction), expanding it in the
absence of 4-OHT for 48 h before labeling the replicating DNA
and performing SMARD. Results indicate that, following 4-OHT
withdrawal, origin activity and the temporal order of replication
revert to pre-induction levels (right portion of Figure 6 and Figure
S6). Therefore, the activity of Pax5 is continuously required to
maintain high rates of origin firing within the CH-39RR region
(these changes are not epigenetically inherited). This also means
that a low firing rate represents the default state for these Igh
origins, an observation that is consistent with the low level of origin
activity observed in all types of non-B cells examined to date [2].
caused by a different mechanism (e.g., transcription, chromatin
remodeling, etc.).
But how is the regulation of the firing rate of origins achieved
during cell differentiation? It is important to note that the population of double-labeled DNA molecules originates from cells
that are in S phase at the time of the label switch, having crossed
the G1/S transition 0 to 809 before each molecule started
replicating (Figure S7). This means that the 7-fold increase in the
firing rate observed after 1 h of induction (Figure 6H) takes place
in cells that were first exposed to 4-OHT at a time in the cell cycle
close to the G1/S transition (between 609 before G1/S and 809
into S phase; Figure S7). We know that, in mammalian cells, the
replicative helicase is loaded between the end of mitosis and early
G1, completing the process of origin licensing and the formation of
pre-replication complexes (pre-RCs) as reviewed in [34]. We can
therefore conclude that Pax5 controls the rate of origin firing, and
the temporal order of replication, by affecting a stage in origin
activation downstream of pre-RC assembly. A corollary to this
conclusion is that pre-RCs can form across the CH-39RR region
even in the absence of 4-OHT. Therefore, pre-RCs can form
efficiently even in genomic regions characterized by a low firing
rate of origins.
Discussion
In this study, we provide a model of origin activation that can
explain multiple parameters of DNA replication initiation,
progression, and termination throughout a significant portion of
the S phase. Parameters, such as the location of initiation events
and fork collisions, the efficiency of origin firing, the number and
distribution of replication forks, the timing of origin firing, and the
temporal order of replication can be quantitatively explained by
the interplay between two independent variables: the speed of
replication forks and the firing rate of origins. Only the latter
varies extensively along the locus representing the dominant
variable. The firing rate of origins is also the main variable that is
being regulated during cell differentiation.
For each individual experiment, both independent variables can
be considered constant in time (an approximation that is
sufficiently accurate to describe the events taking place within
the Igh locus during the first few hours of S phase but does not
exclude the possibility of variations over the entire S phase). Other
aspects of DNA replication behave as dependent variables that do
not require dedicated regulatory pathways. This means that the
extensive literature concerning the regulation of origin efficiency
and the timing of origin firing can be reexamined in terms of
changes in the firing rate of origins. For example, the apparent
discrepancies mentioned in the introduction regarding some
earlier studies [2,3,5] could be reconciled by changes in the firing
rate of origins during cell differentiation. Hence, our model can
provide a simple conceptual framework to explain the dynamics of
DNA replication at a specific mammalian locus.
Our study suggests that potentially active origins are present at
high density throughout the Igh locus in both committed and
uncommitted pro-B cells (Figures 2A, 4A, S2E, S4D). Active
origins were detected even within portions of the locus that
replicate last (e.g., PmeI#4 in Figure 2). This interpretation is
consistent with recent observations showing the presence of active
origins of replication in portions of the human IGH locus
previously thought to contain only silent ones [13,35]. However,
based on the analysis of a larger set of parameters of DNA
replication (representing all stages of replication), we showed that
the initiation events detected experimentally can be explained by
an entirely stochastic mechanisms of activation (meaning that
origin firing can occur in any unreplicated portion of the Igh locus,
at any time during the S phase).
Events Prior to the Assembly of Pre-RCs Have Only Minor
Effects on the Firing Rate of Origins and the Temporal
Order of Replication
Are the effects of Pax5ER induction limited to the late stages of
origin activation? Figure 6H shows that the firing rate of the CH39RR origins changes over time, peaking at 6 h induction and
returning to the level of committed-pro-B cells thereafter (at 12,
24, and 28 h). We know that the cell cycle of KO-Pax5ER pro-B
cells is approximately 14 h and that their G1 lasts about 6 h (A.D.
and P.N., unpublished observation). This means that the 14-fold
increase in firing rate that we observe during the first 6 h of
induction is linked to events taking place during the preceding G1
(mostly after the assembly of the pre-RCs). In contrast, the firing
rate measured at 12 h of induction is the result of events taking
place not only after pre-RC assembly but also before (during the
previous M, G2, and S phases). Since this value is higher than the
pre-induction level but 63% lower than the firing rate measured at
6 h (Figure 6H), we conclude that some inhibition of the firing rate
takes place during the previous M, G2, and S phases. However,
this inhibition is insufficient to fully compensate for the increase in
firing rate that takes place during G1. We conclude that the firing
rate of the CH-39RR origins is affected by the induction of
Pax5ER at various stages of the cell cycle but is dominated by
events that occur after pre-RC assembly.
The Activity of Pax5 Is Continuously Required to Maintain
a High Rate of Origin Firing within the CH-39RR Region
Are the changes in firing rate induced by Pax5 stable? Or do
they need to be reestablished at every cell division? To address this
question, we took advantage of the fact that the activation of
Pax5ER can be slowly reversed by OHT withdrawal. We first
grew KO-Pax5ER pro-B cells in the presence of 4-OHT for 36 h
to induce full B cell commitment and to increase the rate of origin
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Figure 6. Kinetics of the changes in origin activity induced by 4-OHT (and by 4-OHT withdrawal) within the CH-39RR region in KOPax5ER pro-B cells. (A) Labeling scheme utilized for the time-course experiment in asynchronously growing cells. Tr indicates the time interval
during which the double-labeled DNA molecules start replicating (see Table S1 for details). (B–G) Summary of the results obtained by SMARD from
the analysis of 1,013 fully substituted DNA molecules, 818 meeting the standards of measurement, for a total of 202 double-labeled molecules (see
Table S1 for details). The results of the fitting procedure applied to all experimental data sets obtained for the 129/Sv allele are shown as described for
Figure 2. (H) Firing rate of origins calculated from the simultaneous fit of all experimental data sets obtained for the 129/Sv allele, including those
presented in Figure 5 (**).
doi:10.1371/journal.pbio.1001360.g006
promoting chromatin looping within the CH-39RR region [44].
Hence, it seems plausible that chromosomal topology and domain
boundaries play a role in modulating the firing rate of origins
within the Igh locus. In this context, the role of Pax5 could be
helping to form new chromosomal loops or altering the topology
of preexisting ones. Similar mechanisms could provide a way to
affect the activity of multiple origins of replication across large
sections of the genome (perhaps by changing their accessibility to
rate-limiting factors or by facilitating DNA unwinding at origins).
In fact, preliminary evidence indicates that the deletion of short
regulatory elements can affect the firing rate of origins across
entire Igh domains (P.N., unpublished result).
One of the appeals of the model of origin regulation presented
above is that physiological variations in a range of parameters of
DNA replication can automatically be compensated without
necessarily requiring the intervention of checkpoints. For example,
a localized decrease in replication fork progression will provide
more time for origins to fire, resulting in the activation of
additional origins (proportionally to their firing rate). This can
limit the probability that sections of the genome will remain
unreplicated for extended periods of time. A similar compensatory
mechanism can operate at the genomic level as a result of cell-tocell variations in origin licensing, or due to factors that are ratelimiting for origin firing [45]. Even in the presence of a drastic
reduction in the global level of origin activity (e.g., following the
activation of DNA damage checkpoints), residual initiation events
are expected to continue to occur according to the differential in
firing rate of origins in the various portions of the genome
(therefore preserving the temporal order of replication). These
predictions are consistent with experimental observations such as
those referring to the activation of dormant origins during
replicative stress [46,47]. Thus, this model of origin regulation
can help us to explain experimental observations that are not fully
understood.
Our study also suggests a significant degree of coordination in
the level of origin activity across large sections of the Igh locus (tens
to hundreds of kilobases in size). Within these domains, the firing
rate appears to be relatively uniform and is modulated synchronously, over a broad range, as a result of changes in cell
differentiation. The temporal order of replication is determined by
the combined effect of variable domain sizes and firing rates.
Consistent with studies performed at other mammalian loci [26],
we showed that groups of origins can be deleted from the Igh locus
without significantly affecting the firing rate of the remaining
origins, or the location of the firing rate transitions (these two
qualities of chromosomal domains are not significantly affected by
the activity of individual origins of replication). In contrast,
clustering of yeast origins with similar characteristics has been
observed in various studies, but the firing rate of each origin is
different and appears to be individually regulated [15,36,37].
Hence, our results point to significant differences in the way origin
activity is regulated in the two organisms, perhaps reflecting a role
of domain boundaries in the modulation of mammalian DNA
replication (see below) [38].
During B cell commitment, the largest changes in firing rate
occur in Igh domains that contain Pax5 binding sites and depend
on the continuous expression of this protein. This suggests that
cell-specific factors participate in controlling the temporal order of
replication by regulating the firing rate of origins within specific
chromosomal domains. Pax5 modifies the firing rate of origins
using at least two separate mechanisms that operate at different
times during the cell cycle. One mechanism acts upstream of preRC assembly. It produces a stable decrease in the peak rate of
origin firing across the CH-39RR region, suggesting it affects a
limiting step in origin activation (perhaps the formation of preRCs). However, the limited magnitude of the change means that
this mechanism can only marginally affect the temporal order of
replication. In contrast, within the CH-39RR region, the rate of
origin firing is mostly determined by events occurring in late G1/
early S (after the loading of ORC and MCM proteins). This
second mechanism produces a rapid increase in the firing rate of
origins that continues for a few hours after the induction of
Pax5ER (cells need to be exposed to Pax5 for most of G1 in order
to reach the maximum level of origin activity). These results are
consistent with recent observations suggesting that temporal order
of replication is strongly affected by events downstream of the
assembly of pre-RCs [39,40].
Given that Pax5 affects origin activity in multiple ways,
dissecting its activity will require additional studies. However, it
is important to point out that chromosomal loops comparable to
the CH-39RR, DH-JH, proximal-VH, and middle-VH domains
have been recently identified by 4C and 3C assays in C57BL/6
committed pro-B cells [41]. This suggests that the sharp transitions
in firing rate detected in our study may occur at the bases of
chromosomal loops; with plateaus making the body of each loop
(adjacent loops with a similar firing rate may not be distinguishable). Interestingly, the loop organization of the Igh locus can
change during B cell differentiation [42]. In addition, Pax5 has
been shown to alter the chromosomal architecture of various
portions of the locus, inducing locus contraction [43], and
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Materials and Methods
Pro-B Cell Cultures Preparation and Characterization
Short-term bone marrow pro-B cell cultures were prepared as
previously described [48]. Briefly, bone marrow cells were isolated
from an 8-wk-old Rag22/2 mouse (129/Sv) and plated at 106
cells/ml in RPMI supplemented with 5% heat-inactivated fetal
calf serum (FCS; Invitrogen), 2 mM L-glutamine, 16 penicillinstreptomycin, 50 mM 2-mercaptoethanol, and 10 ng/ml IL-7
(R&D). Cells were kept in culture for 7 d before labeling the
replicating DNA with halogenated nucleotides. FACS analysis
performed before DNA labeling indicated that more than 99% of
the cells in culture were CD19+ (corresponding to committed proB cells). In contrast, pro-B cells from Pax52/2 and Pax52/2
Rag22/2 mice can be grown ex vivo for extended periods of time
[49]. Long-term uncommitted pro-B cell cultures were prepared
from the bone marrows of 1-wk-old Pax52/2 Rag22/2 (129/Sv)
and Pax52/2 (129/Sv-C57BL/6) mice, as previously described
[20]. These cells were expanded ex vivo on irradiated feeder cells
in IMDM medium supplemented with 2% heat inactivated FCS,
0.15% primatone, 2 mM L-glutamine, 16penicillin-streptomycin,
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50 mM 2-mercaptoethanol, and 1 ng/ml IL-7. The mouse stromal
cell line ST2 was used to prepare the feeder layers for pro-B cell
cultures. These cells were grown in IMDM medium supplemented
with 10% heat-inactivated FCS, 2 mM L-glutamine, 16 penicillin-streptomycin until 60% confluent, then irradiated (11 Gy) and
used as feeders. All pro-B cell cultures were maintained as
polyclonal populations. However, a clonal population was also
derived from the bone marrow of a Pax52/2 mouse (129/SvC57BL/6) and used for some of the experiments presented in this
study.
The location of Pax5 binding sites in committed pro-B cells, as
well as their DNA sequence, chromatin features, and transcriptional activity, is presented for the entire Igh locus in a previous
publication [28]. Briefly, within the 1.4 Mb region considered in
the current study (from strain 129/Sv), the DNA sequences for six
of these sites are present at 11 different locations (marked from 1
to 11 in Figure 4E). The sites marked from 1 to 4 have unique
DNA sequences that align to the reference mouse genome
(strain C57BL/6) at the following positions: assembly mm8_
chr12:113895500; and chr12:113972530-113972740. In contrast,
sites 5 and 11 have the same DNA sequence and align to the
reference strain at position mm8_chr12: 114630500-114630900,
while sites 6–10 align at position mm8_chr12:114541600114542000. With the exception of site number 4, all other sites
display active chromatin marks in committed pro-B cells (H3K9ac
and H3K4me2 [28]). However, these marks depend on Pax5
binding only at sites 1 and 2. Hence, there is no simple correlation
between the firing rate of origins and the presence of active
chromatin marks. Moreover, unlike Pax5 binding sites located in
the distal portion of the Igh locus, none of these sites is associated
with PAIR elements and Pax5-dependent antisense transcription
[28].
A portion of the uncommitted pro-B cell culture was transduced
with the retroviral vector Pax5ER-IRES-GFP that we previously
used to reconstitute Pax5 expression in uncommitted pro-B cells
[20,30,31,50]. In these experiments, GFP expression allowed us to
identify the cells successfully transduced with the retroviral vector.
The populations of GFP+ cells were isolated by FACS in sterile
conditions using a FACS MoFlo apparatus (sorting buffer: 16
PBS, 4% FBS, 16 penicillin-streptomycin). We have previously
shown that the fusion protein Pax5ER can respond within minutes
of the addition of 4-OHT (1 mM), resulting in the efficient
expression of target genes (usually within a few hours from
induction [30]). Pax5ER expression was monitored by immunoblot using antibodies raised against the amino terminus of Pax5
(Santa Cruz), or against the carboxy terminus of the estrogen
receptor alpha (Santa Cruz), to determine the total level of
expression in 36105 cells, using the expression of beta-actin as a
reference (Santa Cruz). The induction of Pax5ER was monitored
by FACS using anti-mouse-CD19 antibody (Pharmingen). Following 4-OHT induction, CD19+ and CD192 pro-B cells were
isolated using an AutoMACS cell separator and anti-CD19
conjugated magnetic beads (Miltenyi), according to the manufacturer’s procedure. Cell-cycle analysis before and after 4-OHT
induction was performed by FACS following propidium-iodide
staining of the cells (50 mg/L) in hypotonic buffer (0.1% sodium
citrate). The lack of DNA-damage checkpoint activation was
determined on total cell lysates from 36105 cells in the presence of
a protease and phosphatase-inhibitors cocktail (Thermo Scientific),
using antibodies raised against p53 (Cell Signaling), phospho-p53,
Ser15 (Cell Signaling), phospho-histone H2A.X, Ser139 (Cell
Signaling), phospho-Chk1, Ser345 (Cell Signaling), phosphoRad17, Ser645 (Cell Signaling), and phospho-ATM, Ser1981
(Santa Cruz).
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Double-Labeling of Replicating DNA, DNA Purification,
and SMARD
A detailed procedure for SMARD is presented in previous
publications [17,21,22]. Briefly, cells in the exponential phase of
growth were sequentially labeled with IdU and CldU (25 mM each)
for periods of time between 3 and 4 h at a density between 36105
and 86105 cells/ml. The duration of each of the labeling periods
was chosen to be longer than the replication time of the fragments
analyzed but short enough to prevent multiple rounds of replication according to the principles described in previous
publications [17,21]. Total genomic DNA was prepared in
agarose plugs (about 106 cells/plug) and digested with either PacI,
SwaI, or PmeI (40 units/plug). Restriction fragments were purified
by pulsed-field gel electrophoresis, recovered by agarase digestion,
and stretched on silanized microscope slides by capillary action.
During SMARD, the restriction fragments of interest were
detected with the following probes: SwaI #2 was detected with
several PCR products spanning the positions 14,606–25,310 and
35,667–46,206 of the Igh clone BAC199M11 (GeneBank:
AF450245), together with a plasmid clone containing the switch
region c1 (pc1/EH10; [51,52]); PacI #3 was detected with the
plasmids UUGC2M0237A15 and UUGC2M0215F07 (GeneBank: AZ966978, AZ966626/AZ951255, AZ950883), and with
PCR products spanning, respectively, positions 9530909–
9543550, 9560850–9571463, and 9644195–9652184 in the mouse
chromosome 12 contig (GeneBank: NT_039553); PmeI #4 was
detected with PCR products spanning, respectively, positions
1204509–1206646, 1206954–1208600, 1209395–1211337, and
1212606–1213479 in GeneBank: AJ851868; and PmeI #5 was
detected with the plasmids UUGC1M0532J03, UUGC1M0259G14, UUGC2M0265N20, UUGC1M0166E11, and UUGC2M0258M14 (spanning positions 211454–219261, 413828–421730,
591439–598359, 631282–640007, and 647267-663409 in GeneBank: AJ851868). The same probes were used for the detection
of both the 129/Sv and the C57BL/6 alleles. Microscopy image
alignment to the maps of the restriction fragments and data
analysis were performed as previously described [22]. Only
molecules unbroken, fully-substituted with halogenated nucleotides, and sufficiency stretched to be aligned were considered.
Numerical Simulation of the SMARD Data
In order to determine how origin firing is regulated in
mammalian cells, we developed a new mathematical formalism
and a simulation procedure which allow us to rigorously analyze
the experimental results obtained by SMARD. The numerical
simulations used in this study allow us to reproduce SMARD data
sets from any given known replication scenario (i.e., a defined
firing rate and fork speed), making it possible for us to study the
statistics of the experimental data sets. As reported in the Results
section, we performed numerical simulations of all experiments
presented in this study. The simulation procedure is described in
detail in reference [18] but can be summarized as follows:
(1) A replication cycle is simulated using a Monte-Carlo
procedure. Replication forks are stochastically created based
on firing rates similar to the ones presented in Figure 2E.
Then the DNA is bi-directionally replicated from each
origin by forks moving at a constant speed until all forks
coalesce.
(2) Once the full replication cycle is simulated, the labeling
procedure is also mimicked by randomly defining the time
when incorporation of halogenated nucleotides starts relative
to the starting time of the replication cycle.
15
July 2012 | Volume 10 | Issue 7 | e1001360
Developmental Control of Origin Activity
(3) A labeled fragment is generated from the simulated
replication cycle and the labeling timeline. The simulations
give us the positions of the forks as a function of time. The
labeled fragments are created from the fork positions at the
times when nucleotide types are switched.
(4) Steps 1 to 3 are repeated many times (typically 5,000 times),
and the recorded simulated fragments are analyzed just as the
real molecules were. In particular, when the number of
analyzed fragments equals that of the actual experiments, we
can use the simulation-to-simulation variations of measured
quantities to estimate errors of those quantities. This leads to
the error bars shown in various figures, such as Figure 2B–C
(fork densities), Figure 2D (IdU content), and Figure S2B
(replication time).
where x1 and x2 are the boundaries of the region of interest
(Equation 3). This calculation approach provides us with a faster
and more precise way to describe the average replication kinetics
for a specific genomic region, and its results were validated using
simulations of known replication scenarios, as described in the
previous section. In addition, simulations are still needed in order
to study the statistical fluctuations of f(x, t) and r6(x, t).
Computer Modeling of the SMARD Data
Fitting Experimental Data
average number of initiation events occurring within a specific
section of the locus, per allele, during each S phase as:
Ni ~
0 x1
Iðx,tÞ½1{f ðx,tÞ$dxdt,
This calculation technique was then used to fit the experimental
data sets to parameterized functional forms for I(x, t) and v6(x, t).
This means that starting with a series of parameters defining both
the firing and the velocity profiles, we used our calculation
technique to produce the corresponding SMARD-like results. The
parameters were then iteratively adjusted in order to obtain the
best fit between the calculated values and the experimental
measurements. For the fits performed in this study, we assumed
homogeneous fork speeds throughout both space and time (i.e.,
v6(x, t) = v for left- and right-moving forks). We also assumed that
the firing rate is constant throughout time but spatially inhomogeneous, I(x, t) = I(x). A larger firing rate implies a shorter average
replication time and vice versa. Time-dependent firing rates
were considered but were not needed to obtain a satisfactory
representation of the data. In other words, at the hundredkilobase-to-megabase scale considered in this study, the replication
timing is dominated by spatial variations in the firing rate. Our
calculation technique also provided us the distribution of times
required to duplicate a given section of the genome.
The SMARD data used in the fit are shown as empty symbols
in Figures 2B–D and S2B. For each experiment, we performed a
global fit of the two replication fork distributions (solid lines in
Figure 2B–C) of the IdU content (solid lines in Figure 2D), and
of the replication times for all restriction fragments (filled
symbols in Figure S2B). Error bars on the experimental data
were obtained from numerical simulations where each simulated
experiment uses the best-fit parameters to collect the same
number of labeled DNA fragments analyzed in the real
experiment. Figure S2D shows the free parameters obtained
from the fits, with the firing rate of origins I(x) also graphically
illustrated in Figure 2E. For the purpose of the calculations, the
firing rate was considered to be composed by a constant
background of initiation events (one parameter), to which
additional events are added where initiations are more likely to
occur (active origins clusters). For simplicity, these initiation
clusters were assumed to have the shape of a rounded box
defined by three parameters (position and width of the cluster,
and the firing rate within the cluster). While this is likely to
represent an oversimplification of the conditions existing in vivo,
we found that more complex shapes were not required to obtain
meaningful representations of the experimental data sets. In
contrast, Gaussian curves were tested but resulted in fits that do
not accurately reproduce the observed number of initiation
events and their location. Hence, curves where the firing rate
changes sharply along the genome provide a better representation of the experimental data sets than curves where the firing
rate changes gradually. In Figure 2E, two additional parameters
In principle, numerical simulations could also be used to
determine the most likely set of parameters responsible for
generating specific experimental data sets obtained by SMARD.
However, the need to explore a large number of variables makes
this fitting procedure impractical and time consuming when the
replication scenario is not pre-determined. Therefore, we also
developed an analytical formalism that allows us to rapidly model
the results obtained by SMARD [18]. The formalism leads to rate
equations that describe changes in the number of replication forks
as a function of both the location along the genome (x) and the
time elapsed since the beginning of S phase (t). More specifically,
we define a set of three coupled partial differential equations that
describe the space-time evolution of the replication fraction, f(x, t),
and the two densities of forks, r6(x, t), moving rightward (+) and
leftward (–). Equation 1 gives the rate of change of the probability
that a specific location x along the genome is replicated at time t,:
Lf ðx,tÞ
~vz ðx,tÞrz ðx,tÞzv{ ðx,tÞr{ ðx,tÞ,
Lt
where v6(x, t) is the speed of the replication fork at the position x at
time t. This first equation states that the rate of replication of a
specific location is proportional to the number of forks at the same
location times their respective replication time. The rate of change
of the fork densities (i.e., the number of forks moving in a given
direction per kb) is given by Equation 2:
!
"
Lr+ (x,t) L v+ ðx,tÞr+ ðx,tÞ
~Iðx,tÞ½1{f ðx,tÞ${
+
Lx
Lt
½vz ðx,tÞzv{ ðx,tÞ$rz ðx,tÞr{ ðx,tÞ
,
1{f ðx,tÞ
where I(x, t) is the firing rate that corresponds to the number of
initiation events per time per unreplicated DNA length. The two
terms on the right-hand side of the latter equation represent
initiation and termination events, respectively, while the terms on
the left-hand side express the space-time-propagation of the
replication forks. This second equation defines the rate of change
of the probability of observing a replication fork at a specific
location and time in the form of a classic transport equation. If
both the firing rate and the fork speed space-time functions are
known, these equations can be numerically integrated to obtain
f(x, t) and r6(x, t). Once our set of equations is solved, the results
can be converted to profiles of IdU content and fork distributions,
as they are observed in the SMARD experiment (for details, see
reference [18]). The solution can also be used to calculate the
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?
ð xð2
16
July 2012 | Volume 10 | Issue 7 | e1001360
Developmental Control of Origin Activity
Figure S2 Replication of the Igh locus in Pax52/2 Rag22/2
show the level of origin firing outside the genomic region
investigated. The value of these parameters may be seen as an
effective constant firing rate profile outside the studied region.
Finally, one parameter was used to fit the velocity of fork
propagation (meaning that the speed of replication forks was
assumed to be constant throughout the region analyzed).
Among the other results of the fit, Figure S2F shows the probability for each section of the genome to be replicated by forks
that originated within the modeled region, while Figure S2G
shows the probability of being replicated by replication forks
moving leftward or rightward. Finally, Figure 2F shows the
average replication kinetics of the modeled genomic region. The
color value on this space-time graph indicates the probability for
each genomic position to replicate at a specific time in S phase.
The transition from unreplicated (orange) to replicated (blue)
occurs at a different speed in a different portion of the genome.
Three contour lines are shown to illustrate which regions
replicate faster than others (showing when 10%, 50%, and 90%
of cells have replicated the corresponding portion of the
genome).
The images collected by SMARD represent all stages of DNA
replication. As a result, the number of initiation and termination
events scored on individual DNA molecules (Table S1, columns g
and h) is lower than the actual number of events taking place
during every replication cycle. For example, initiation events can
be easily scored when the size of the IdU patch is small, but no
longer so at more advanced stages of replication (when one of the
outward-moving forks runs off the fragment or coalesces with
another fork). In contrast, our calculation method determines the
actual number of initiation events and fork collisions occurring, on
average, every replication cycle. Hence, in order to compare the
outcomes of these two procedures, we performed additional
simulations of the SMARD experiments to determine how many
of initiations/collisions would be observed experimentally from the
results of each fit. Figure S2C shows that the number of events
obtained from the fit is comparable to the number observed
experimentally.
uncommitted pro-B cells (129/Sv). (A) Image of a double-labeled
DNA molecule containing two initiation events and two externally
generated replication forks. (B) Graphical representation of the
replication times (Tr) for each restriction fragment obtained
experimentally by SMARD (gray; see Table S1 for details) and
from the fit (red). Error bars were calculated as described in
Materials and Methods. (C) Numbers and percentages of initiation
events and fork collisions detected in the population of doublelabeled DNA molecules by SMARD, or calculated from the fit. (D)
Miscellaneous results from the fit including the most likely size and
position of the active origin clusters. (E) Schematics of the four
restriction fragments analyzed in this experiment (shown to scale).
Black ovals indicate the position of the midpoints for the initiation
events detected in these experiments (where the corresponding
origins are more likely to be located). Multiple ovals overlapping
the same portion of the genome are a hallmark of origin mapping
saturation (it indicates that the same origin has been mapped on
different DNA molecules). The rarity of these overlaps indicates
that origin density is higher than depicted. (F) Probability for
various portions of the locus to be replicated by origins located
within the genomic region under investigation (yellow), or near the
distal-VH genes (red), or downstream of the Igh locus (blue), as
calculated from the fit. (G) Probability for various portions of the
locus of being replicated by forks moving rightward (red) or
leftward (blue), as calculated from the fit. Note that the probability
changes in relation to the firing rate of origins (gray dotted line).
(TIF)
Figure S3 Replication of the Igh locus in a DH-JH rearranged
clonal population of Pax52/2 uncommitted pro-B cells (129/SvC57BL/6). The results for each allele are shown separately. (A–F)
Summary of some of the results obtained by SMARD and from
the fitting procedure (as described for Figure S2).
(TIF)
Figure S4 Replication of the Igh locus in Rag22/2 committed
pro-B cells (129/Sv). (A–F) Summary of some of the results
obtained by SMARD and from the fitting procedure (as described
for Figure S2). For three of the restriction fragments analyzed in
these cells, the number of double-labeled DNA molecules
displayed the occurrence of more than one initiation event is
significantly higher than in uncommitted pro-B cells (9%, 14%,
and 12%, respectively, for SwaI #2, PmeI #4, and PmeI #5). This
is consistent with an increase in the average number of origins
firing within the locus during each replication cycle (see main text).
(TIF)
Supporting Information
Figure S1
Single molecule analysis of replicated DNA. DNA
labeling scheme used for SMARD. Within a population of
exponentially growing cells, DNA synthesis is limited to cells
transiting through the S phase. If a specific portion of the cellular
genome is considered (e.g., a restriction fragment, gray bars), the
distribution of replication forks within this region will be
representative of all stages of DNA replication (steady state
distribution). Following a first labeling period of 3 h with IdU
(central panel), each cell will be at a different position in the cell
cycle and the halogenated nucleotide will be distributed among the
DNA molecules as indicated by the red color. Only the DNA
molecules that started and completed their replication during the
labeling period will be fully substituted with IdU, while the others
will be either partially substituted or not labeled. Following a
second labeling period of 3 h with CldU (right panel), halogenated
nucleotides will be distributed among the DNA molecules as
indicated by the red and green colors (IdU and CldU,
respectively). During SMARD, only the molecules fully substituted
with one or both nucleotide analogs are considered. Within the
population of double-labeled DNA molecules, the distribution of
IdU-CldU transitions corresponds to the distribution of replication
forks at the time of the label switch.
(TIF)
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Figure S5 Reconstitution of Pax5 expression in Pax52/2
uncommitted pro-B cells (129/Sv-C57BL/6). (A) Level of
expression of Pax5ER determined using a monoclonal antibody
that recognizes the amino-terminal portion of Pax5. Before 4OHT induction, the level of Pax5ER in KO-Pax5ER pro-B cells is
approximately 20% of the level of wt Pax5 present in the pro-B cell
line 6312. (B) Following 4-OHT induction, the expression of
Pax5ER increases several times, to reach levels comparable to wt
Pax5. (C) Kinetics of expression of the surface marker CD19
following 4-OHT induction. A low percentage of CD19+ cells are
present before the addition of 4-OHT, but this value increases
more than 20 times after the induction of Pax5ER. A small portion
of the KO-Pax5ER pro-B cells remains CD192 even after
prolonged 4-OHT induction. (D–I) Summary of some of the
results obtained by SMARD and from the fitting procedure (as
described for Figure S2). (L) Cell cycle profiles of KO-Pax5ER
pro-B cells before (0 h) and after 4-OHT induction (3 h and 24 h).
These cells continue to cycle even after 4-OHT treatments longer
17
July 2012 | Volume 10 | Issue 7 | e1001360
Developmental Control of Origin Activity
than 1 wk (not shown). (M) Absence of markers of DNA damage
and checkpoints’activation following 4-OHT induction. Immunoblots from two different reconstitution experiments (Pax52/2 and
Pax52/2 Rag22/2) are shown before (2) and after (+) a 3 h
treatment with 1 mM 4-OHT. Similar results were obtained using
induction times between 1 and 24 h, and with antibodies against
phospho-ATM Ser1981, and phospho-Rad17 Ser645 (not shown).
The positive controls for each immunoblot (E) were prepared by
treating KO-Pax5ER pro-B cells with 25 mM etoposide for 2 h.
(TIF)
cases, the Igh alleles were analyzed independently and only the
corresponding mouse strain is indicated. DDH-JH indicates that the
Igh allele was DH-JH rearranged. Column b, duration of the first
and second labeling periods with IdU and CldU (respectively, Tp1
and Tp2). SMARD requires that the duration of each labeling
period is longer than the average replication time of the restriction
fragments under investigation [17,22]. Column c, number of Igh
molecules surveyed by fluorescence microscopy. This number
includes all the molecules detected during each experiment based
on the presence of the hybridization signals and regardless of the
level of substitution with halogenated nucleotides (e.g., fully,
partially, or not substituted). In some of the experiments only the
molecules fully substituted with halogenated nucleotides were
counted; therefore, this value was not determined (n.d.). Columns
d and e, respectively, percentage and number of surveyed
molecules containing one or both halogenated nucleotides. Only
some of these molecules fulfilled the criteria required by SMARD
and were included in column f (see below). Column f, NR, NG, and
NRG indicate the number of imaged molecules substituted with,
respectively, IdU (detected in red), CldU (detected in green), and
both halogenated nucleotides (double-labeled, both red and
green). These numbers include only the molecules suitable for
measurement (fully substituted, unbroken, sufficiently stretched to
be unequivocally aligned to the map of the region, and not
overlapping with other molecules). Column g, number of initiation
events normalized for 100 double-labeled DNA molecules (in red)
and percentage of double-labeled DNA molecules containing
initiation events (in black). These values may differ for some of the
restriction fragments because a fraction of the DNA molecules can
contain more than one initiation event. Column h, number of
colliding forks (termination events) normalized for 100 doublelabeled DNA molecules (in red) and percentage of double-labeled
DNA molecules containing colliding forks (in black). Column i,
average time required to replicate the genomic region encompassed by the restriction fragment. This value (Tr) was calculated
using the following equation: Tr = Tp2 6 NRG/(NG+NRG). A
detailed description of the procedure is provided in previous
publications [17,22]. Column l, average number of replication
forks within the population of double-labeled DNA molecules.
Column m, average speed of replication forks within the restriction
fragment. This value is obtained by dividing the size of the
restriction fragment (from column n) by Tr (from column i) and by
the average number of replication forks (from column l). The
values indicated are expressed in kilobases per minute. Column n,
size of the restriction fragment in kilobases.
(PDF)
Figure S6 Kinetic of the changes in origin activity induced by 4OHT (and by 4-OHT withdrawal) within the CH-39RR region in
KO-Pax5ER pro-B cells. (A–F) Summary of some of the results
obtained by SMARD and from the fitting procedure (as described
for Figure S2). All experimental data sets obtained for the 129/Sv
allele were fitted simultaneously as described in Materials and
Methods. Panel C also shows the positions of the initiation and
termination events detected by SMARD, with numerals indicating
the normalized frequency of the events as described for Figure 2.
(TIF)
Figure S7 Labeling of DNA molecules from fragment SwaI#2
during SMARD. Results presents in Table 1 indicate that SwaI#2
requires approximately 609to complete its replication. At the top of
the picture, this window of time is presented as a gray doubleheaded arrow. In an asynchronous population of cells, this window
of time will be variably distributed in relation to the labeling
periods with IdU and CldU (see double-headed arrows labeled
from1 to 36). This will result in molecules that are variably
substituted with the nucleotide (unlabelled if the molecule started
replicating before time a, partially IdU-labeled if the molecule
started replicating between time a and b, fully IdU-labeled between
time b and c, double-labeled between time c and d, and so on). For
each cell, the window always encompasses a portion of S phase.
From Figures, 2F, 4F, and 5H we know that the replication of
fragment SwaI#2 is complete in more than 90% of cells by 1409
minutes into S phase. Since the fragment takes approximately 609
to replicate, we conclude that the replication of each individual
molecule starts between 09 and 809 (140 – 60 = 80) into S phase
(meaning that the G1/S occurs from 09 to 809 before each
fragment starts replicating). Black double-headed arrows indicate
the possible location of the G1/S transition for the cells that
produce the population of double labeled molecules. We can see
that when 4-OHT is provided 1 h before the label switch a few
cells will have already passed the G1/S transition (e.g., #16), while
many others will have a similar probability of being at the end of
G1 or in early S (e.g., #19–22).
(TIF)
Author Contributions
Table S1 Quantitative analysis of the SMARD data. Column a,
genetic background, cell type and treatment, restriction fragment
name, and mouse strain(s) of origin for the Igh loci analyzed in
each experiment. The name of the mouse strain repeated twice
indicates that the cells used for the experiment were homozygous
and that the two Igh alleles were analyzed together. In all other
The author(s) have made the following declarations about their
contributions: Conceived and designed the experiments: PN. Performed
the experiments: PN AD IV. Analyzed the data: PN MGG JB. Contributed
reagents/materials/analysis tools: MB SK CLS. Wrote the paper: PN, with
limited contributions by JB and CS.
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19
July 2012 | Volume 10 | Issue 7 | e1001360
SUPPLEMENTARY TABLE 1
(a)
(b)
(c)
(d)
(e)
(f)
(g)
(h)
(i)
(l)
(m)
(n)
Cell Type &
Restriction
Fragment
Labeling
time in
(minutes)
Total # of
Igh
molecules
surveyed
% of Igh
molecules
containing
halogenated
nucleotides
Total # of
imaged
molecules
fully
substituted
with IdU,
CldU, or
both
# of Igh
molecules
suitable for
SMARD
# of
colliding
forks
per 100
molecules
% of
molecules
showing
colliding
forks
within
the
fragment
Average time
required to
replicate the
genomic
region
encompassed
by the
restriction
fragment
Tr
Average #
of
replication
forks per
fragment
Average
speed per
replication
fork
(kb/min.)
Fragment
Size
NR_NG_NRG
# of
initiation
events
per 100
molecules
% of
molecules
showing
initiation
events
within
the
fragment
311
70__103___36
0
39
58
1.39
2.25
181
0%
39%
54
1.51
2.24
183
107
1.74
2.98
~555
130
1.68
2.25
~492
142
1
1.20
181
68
1
1.69
~118
111
1.11
1.31
~168
61
1.14
1.15
~141
56
1.74
1.86
181
70
1.39
1.88
183
122
2.14
2.13
~555
99
2.20
2.26
~492
IdU__CldU
Tp1___Tp2
(kb)
Pax5-/Pro-B cells
Pax5-/- Rag2-/Bone Marrow
225___225
942
33%
SwaI
(129/Sv_129/Sv)
Pax5-/- Rag2-/Bone Marrow
225___225
1809
33%
597
119__182__57
51
0
51%
0%
PacI
(129/Sv_129/Sv)
Pax5-/- Rag2-/Bone Marrow
225___225
n.d.
n.d.
242
39___30___27
15
59
7%
48%
PmeI-#4
(129/Sv_129/Sv)
Pax5-/- Rag2-/Bone Marrow
225___225
n.d.
n.d.
154
35___28___38
50
18
47%
18%
PmeI-#5
(129/Sv_129/Sv)
Pax5-/(B.M. clone)
240___240
298
39%
116
28___16___23
0
0
0%
0%
SwaI
(129/Sv, D-J)
Pax5-/(B.M. clone)
240___240
288
40%
115
41___28___11
0
0
0%
0%
PacI
(129/Sv, D-J)
Pax5-/(B.M. clone)
240___240
316
37%
117
26___22___19
0
11
0%
11%
SwaI
(C57BL/6, D-J-r)
Pax5-/(B.M. clone)
240___240
541
34%
184
44___41___14
14
0
14%
0%
PacI
(C57BL/6, D-J-r)
wt Pro-B cells
Rag2-/Bone Marrow
180___180
n.d.
n.d.
227
58___75___34
50
24
41%
24%
SwaI
(129/Sv_129/Sv)
Rag2-/Bone Marrow
180___180
n.d.
n.d.
302
70__104___66
20
20
20%
20%
PacI
(129/Sv_129/Sv)
Rag2-/Bone Marrow
180___180
n.d.
n.d.
234
25___21___44
59
55
45%
36%
PmeI-#4
(129/Sv_129/Sv)
Rag2-/Bone Marrow
180___180
n.d.
n.d.
210
61___56___69
PmeI-#5
(129/Sv_129/Sv)
1
65
55
55%
42%
Pax5ER
Reconstituted
Pro-B cells
KO-Pax5ER
220___220
774
46%
356
97__115___71
w/o 4-OHT
14
18
11%
17%
84
1.32
1.56
173
64
1.33
2.13
181
84
1.64
1.26
173
80
1.73
1.31
181
72
1.79
1.40
181
93
1.15
1.69
181
67
1.29
2.09
181
51
1.38
2.57
181
71
1.81
1.41
181
63
1.97
1.46
181
83
1.88
1.16
181
62
1.78
1.64
181
SwaI
(C57BL/6)
KO-Pax5ER
220___220
808
39%
315
98__113___46
w/o 4-OHT
17
15
15%
13%
SwaI
(129/Sv)
KO-Pax5ER
220___220
n.d.
n.d.
374
102__123__76
28h 4-OHT
CD19+
36
30
33%
26%
SwaI
(C57BL/6)
KO-Pax5ER
220___220
882
49%
432
98__139___80
28h 4-OHT
CD19+
40
33
34%
28%
SwaI
(129/Sv)
KO-Pax5ER
220___220
n.d.
n.d.
355
84__137___66
28h 4-OHT
CD19-
53
26
44%
23%
SwaI
(129/Sv)
Pax5-/- Rag2-/Bone Marrow
180___180
363
27%
98
18___24___26
0
15
0%
15%
36h 4-OHT
(mock induc.)
SwaI
(129/Sv_129/Sv)
KO-Pax5ER
180___180
800
38%
304
81___97___58
36h 4-OHT
CD19+,
16
14
14%
12%
48h w/o 4-OHT
SwaI
(129/Sv)
KO-Pax5ER
180___180
633
18%
114
34___41___16
w/o 4-OHT
19
25
13%
25%
SwaI
(129/Sv)
KO-Pax5ER
180___180
787
15%
118
33___32___21
1h 4-OHT
48
33
38%
33%
SwaI
(129/Sv)
KO-Pax5ER
180___180
1131
13%
147
29___60___32
6h 4-OHT
56
41
56%
38%
SwaI
(129/Sv)
KO-Pax5ER
180___180
453
17%
77
20___28___24
12h 4-OHT
58
29
54%
29%
SwaI
(129/Sv)
KO-Pax5ER
180___180
1150
22%
253
64___97___51
24h 4-OHT
SwaI
(129/Sv)
2
45
33
39%
29%